list of abstracts sesimbra formatted -...
TRANSCRIPT
Bionanotechnology -‐ Recent Advances
10-‐13 July in Sesimbra, Portugal
A satellite meeting to the 9th European Biophysics Congress EBSA 2013
Programme Wednesday 10th July 14.00 -‐ 18.00 Registration 18.00 -‐ 18.10 Introduction 18.00 -‐ 19.00 Workshop Lecture 1: Anthony Watts Nano and sub-‐nano scale distances and dynamics from Solid State NMR in biology – an Introduction 19.00 -‐ 20.00 Dinner and Welcome Reception at the hotel Thursday 11th July 9.30 -‐ 10.30 Workshop Lecture 2: Justin Benesch Mass spectrometry -‐ weighing the evidence for protein structure and dynamics 10.30 -‐ 11.00 Coffee break Research Lectures 11.00 -‐ 11.20 Elizabeth Drolle 11.20 -‐ 11.40 Ilja Kusters 11.40 -‐ 12.00 Joachim Piguet 12:00 -‐ 13.00 Workshop Lecture 3: Peter Hinterdorfer Dynamics of Molecular Recognition: What can we learn from Force Microscopy/Spectroscopy? 13.00 -‐ 14.30 Lunch 14.30 -‐ 15.30 Workshop Lecture 4: Michael Mayer Single Protein Characterization Methods with Nanopores 15.30 -‐ 17.00 Tea break Research Lectures 17.00 -‐ 17.20 Sandra Posch 17.20 -‐ 17.40 Melanie Koehler 17.40 -‐ 18.00 Giovanni Maglia Evening free for participants to make their own dinner arrangements. Friday 12th July 9.30 -‐ 10.30 Workshop Lecture 5: Horst Vogel Analysis of complex transmembrane signaling networks: from single cells to single molecules 10.30 -‐ 11.00 Coffee break Research Lectures 11.00 -‐ 11.20 Fabiola Gutierrez 11.20 -‐ 11.40 Anna Wypijewska 11.40 -‐ 12.00 Patricia Dijkman 12.00 -‐ 13.00 Workshop Lecture 6: David Klenerman Principles of single molecule fluorescence and its application to biology and biomedicine 13.00 -‐ 14.30 Lunch 14.30 -‐ 15.30 Workshop Lecture 7: Dimitrios StamouSingle Small Unilamellar Vesicles: tools for biophysics and technology 15.30 -‐ 17.00 Tea break Research Lectures 17.00 -‐ 17.20 Kata Hajdu 17.20 -‐ 17.40 Krisztina Nagy 17.40 -‐ 18.00 Raul Pacheco-‐Gomez Workshop drinks and dinner, Restaurant "O Canhão", next to the hotel Saturday 13th July 9.30 -‐ 10.30 Workshop Lecture 8: Helmut Grubmueller Forces and Conformational Dynamics in Biological Nanomachines 10.30 -‐ 11.00 Coffee break Research Lectures 11.00 -‐ 11.20 Francesco Colizzi 11.20 -‐ 11.40 Melinda Magyar 11.40 -‐ 12.00 Liliana Lighezan 12.30 -‐ 13.30 Lunch 13.30 Leave for Lisbon
Travel Travel to Sesimbra Get the metro at Lisbon Airport to Praça Espanha station (www.metrolisboa.pt/) At Praça de Espanha there are coaches going to Sesimbra. The coach company is “TST Transportes Sul do Tejo” bus number 207 (http://www.tsuldotejo.pt/?idioma=2) A timetable can be found below.
Travel from Sesimbra to Lisbon Travel from this meeting to the main congress venue will be organised for Saturday afternoon, after lunch in Sesimbra. Sesimbra Hotel & SPA Address: Rua Navegador Rodrigues Soromenho 2970-‐773 Sesimbra PORTUGAL Tel. geral +351 212 289 800 Tel. reservas +351 212 289 803 Bus 7 Schedule (Lisbon to Sesimbra):
Mass spectrometry -‐ weighing the evidence for protein structure and dynamics
Justin Benesch
Chemistry Department, University of Oxford, UK Mass spectrometry is a recognised approach for characterising proteins and the complexes they assemble into. This application of a long-‐established physico-‐chemical tool to the frontiers of structural biology has stemmed from experiments performed in the early 1990s. While initial studies focused on the elucidation of stoichiometry by means of simple mass determination, developments in mass spectrometry technology and methodology now allow researchers to address questions of shape, inter-‐subunit connectivity, and protein dynamics. Here we will discuss the remarkable rise of mass spectrometry and its application to biomolecular complexes over the last two decades.
Forces and Conformational Dynamics in Biomolecular Nanomachines Helmut Grubmueller
Max-‐Planck Institute for Biophysical Chemsitry, Gottingen, Germany
Proteins are biological nanomachines. Virtually every function in the cell is carried out by proteins -‐-‐ ranging from protein synthesis, ATP synthesis, molecular binding and recognition, selective transport, sensor functions, mechanical stability, and many more. The combined interdisciplinary efforts of the past years have revealed how many of these functions are effected on the molecular level [1]. Computer simulations of the atomistic dynamics play a pivotal role in this enterprise, as they offer both unparalleled temporal and spatial resolution. With state of the art examples, this talk will explain the basics of this high performance computing method [2], the type of questions that can (and cannot) be addressed, and its (current) limitations. The examples include aquaporin selectivity [3,4], mechanics of energy conversion in F-‐ATP synthase [5,6], the mechanical properties of viral capsids [7,8], and tRNA translocation within the ribosome. We will further demonstrate how atomistic simulations enable one to mimic, one-‐to-‐one, single molecule experiments such as FRET distance measurements, and thereby to enhance their accuracy [9,10]. We will, finally, take a more global view on the 'universe' of protein dynamics motion patterns and demonstrate that a systematic coverage of this 'dynasome' allows to predict protein function more reliably [11]. [1] Puchner EM, Alexandrovich A, Kho AL, Hensen U, Schafer LV, Brandmeier B, Grater F, Grubmuller H, Gaub HE, Gautel M. Mechanoenzymatics of titin kinase. PNAS 105: 13385-‐13390 (2008) [2] Hess B, Kutzner C, van der Spoel D, Lindahl E. GROMACS 4: Algorithms for Highly Efficient, Load-‐Balanced, and Scalable Molecular Simulation. J. Chem. Theory Comput. 4: 435-‐447 (2008) [3] de Groot BL, Grubmüller H. Water permeation across biological membranes: Mechanism and dynamics of Aquaporin-‐1 and GlpF. Science 294: 2353-‐2357 (2001) [4] de Groot BL, Grubmüller H. The dynamics and energetics of water permeation and proton exclusion in aquaporins. Curr. Opin. Struct. Biol. 15: 176-‐183 (2005) [5] Böckmann R, Grubmüller H. Nanoseconds molecular dynamics simulation of primary mechanical energy transfer steps in F1-‐ATP synthase. Nature Struct. Biol. 9: 198-‐202 (2002) [6] Czub J and Grubmüller H. Torsional elasticity and energetics of F1-‐ATPase. PNAS 108; 7408-‐7413 (2011) [7] Zink M and Grubmüller H. Primary changes of the mechanical properties of Southern Bean Mosaic Virus upon calcium removal. Biophys. J. 98: 687-‐695 (2010) [8] Zink M, Grubmüller H. Mechanical properties of the icosahedral shell of southern bean mosaic virus: a molecular dynamics study. Biophys J. 96(4): 1350-‐63 (2009) [9] Wozniak AK, Schroder GF, Grubmuller H, Seidel CA, Oesterhelt F. Single-‐molecule FRET measures bends and kinks in DNA. Proc. Natl. Acad. Sci. USA 105(47): 18337-‐42 (2008) [10] Hoefling M, Lima N, Haenni D, Seidel CAM, Schuler B, Grubmüller H. Structural Heterogeneity and Quantitative FRET Efficiency Distributions of Polyprolines through a Hybrid Atomistic Simulation and Monte Carlo Approach. PLOS ONE 6: e19791 (2011) [11] Hensen U, Meyer T, Haas J, Rex R, Vriend G, Grubmüller H. Exploring Protein Dynamics Space: The Dynasome as the Missing Link between Protein Structure and Function. PLoS ONE 7, e33931 (2012)
Dynamics of Molecular Recognition: What can we learn from Force
Microscopy/Spectroscopy?
Peter Hinterdorfer
Institute for Biophysics, Johannes Kepler University Linz, Gruberstr. 40, A-‐4020 Linz,
Austria
In single molecule recognition force spectroscopy (SMRFS), ligands are covalently attached to atomic force microscopy tips for the molecular recognition of their cognitive receptors on probe surfaces. A ligand-‐containing tip is approached towards the receptors on the probe surface, which possibly leads to formation of a receptor-‐ligand bond. The tip is subsequently retracted until the bond breaks at a certain force (unbinding force). Varying the dynamics of the experiment reveals the dependence of the unbinding force from the loading rate. These studies give insight into the molecular dynamics of the receptor-‐ligand recognition process and yield information about the binding pocket, binding energy barriers, and kinetic reaction rates. Applications on isolated proteins, native membranes, viruses, and cells will be presented. We have also developed a method for the localization of specific binding sites and epitopes with nm positional accuracy, termed topography and recognition (TREC) imaging. A magnetically driven AFM tip containing a ligand covalently bound via a tether molecule is oscillated at a few nm amplitude while scanning along the surface. In this way, topography and recognition images are obtained simultaneously. Finally, we will show how high-‐speed bio-‐AFM is able to film the dynamics of recognition processes on the nano-‐scale.
Principles of single molecule fluorescence and its application to biology and biomedicine
Dave Klenerman
Department of Chemistry, Cambridge University Single molecule fluorescence has become a powerful biophysical tool to detect rare species and probe the structure and dynamics of biomolecules in solution, on surfaces and on and in live cells over the last 15 years. This work has encouraged the active collaboration of physical scientists and biologists leading to the development of new and enabling methodologies and providing new insights into fundamental biological processes. I will briefly review the history of the field and then describe the key single molecule fluorescence methods that are currently available to researchers and their principles of operation. Next some of the landmark experiments over the past 15 years will be described, illustrating how the methodology has evolved and that problems of increasing biological complexity can now be tackled. Lastly I will show how this approach can lead to the development of new and powerful technologies by describing the work that lead to the development of the Solexa next generation DNA sequencing method.
Single Protein Characterization Methods with Nanopores
Michael Mayer Department of Biomedical Engineering and Department of Chemical Engineering
University of Michigan, Ann Arbor, MI, USA
Synthetic and biological nanopores can be used for fundamental and applied studies of individual biomolecules in high throughput. By measuring resistive current pulses during the translocation of single molecules through an electrolyte-‐filled nanopore, this technique can characterize the size, conformation, assembly, and activity of hundreds of unlabeled molecules within seconds. Inspired by the olfactory sensilla of insect antennae, we demonstrate that coating nanopores with a fluid lipid bilayer considerably extends the capabilities of nanopore-‐based assays. For instance, coating nanopores with different lipids allows fine control of the surface chemistry and diameter of nanopores. Incorporation of mobile ligands in the lipid bilayer imparts specificity to the nanopore for targeting proteins and introduces control of translocation times for targeted proteins based on their net electric charge. Most recently, we explored the potential of this technique for determining the affinity constant of a protein-‐ligand interaction, monitoring the kinetics of binding of this interaction, characterizing the aggregation state of Alzheimer’s disease-‐related amyloid peptides, as well as determining the molecular shape, dipole moment and rotational diffusion constant of individual proteins.
Single Small Unilamellar Vesicles: tools for biophysics and technology
Dimitrios Stamou
Nano-‐Science Center, University of Copenhagen, Denmark
The functional nanoscale architectures of cells are made of lipid and proteins interacting in a highly coordinated fashion. We are interested in elucidating mechanisms driving nanoscale membrane rearrangements and also in using this knowledge to create functional artificial biomimetic architectures for synthetic biology applications. Here I will discuss the use of single small unilamellar vesicles as versatile tools that can be used both in biophysics and synthetic biology.
Analysis of complex transmembrane signaling networks: from single cells to single molecules
Horst Vogel
Ecole Polytechnique Fédérale de Lausanne (EPFL), Lausanne, Switzerland
G-‐protein-‐coupled receptors (GPCRs) are ubiquitous mediators of signal transduction across cell membranes and constitute a very important class of therapeutic targets. I will present two bioanalytical platforms to control, manipulate and investigate molecular events in GPCR-‐mediated signaling network at a nanometer and attoliter scale.
The first part concerns an approach enabling the study of GPCRs in their native membrane transferred inside-‐out from live cells to lectin-‐coated beads [1]. The access to both sides of the plasma membrane and the receptor allows controlled supply of fluorescent extracellular ligands and intracellular G proteins. Here, the interactions between the different signaling partners during the formation and subsequent dissociation of the ternary signaling complex on single beads can be observed in real time using multicolor fluorescence microscopy. This method of tethering native cellular membranes from live cells and access them from both sides represent a generic platform for investigating complex signaling processes at plasma membranes at the sub-‐micrometer scale.
The second part reports on the investigation of GPCR signalling in single, submicrometer-‐sized native vesicles, derived from living mammalian cells using chemicals or optical tweezers [2, 3]. The vesicles comprise parts of a cell’s plasma membrane and cytosol and represent the smallest autonomous containers performing cellular signaling reactions, thus functioning like minimized cells. Using fluorescence microscopies, we measured in individual vesicles the different steps of GPCR signaling like ligand binding to receptors, subsequent G-‐protein activation and finally receptor deactivation via arrestin translocation. Observing cellular signaling reactions in individual vesicles opens the door for downscaling bioanalysis of cellular functions to the attoliter range, multiplexing single cell analysis, and investigating receptor mediated signaling in multiarray format.
[1] S Roizard, C Danelon, G Hassaïne, J Piguet, K Schulze, R Hovius, R Tampé, H Vogel: Activation of G-‐Protein-‐Coupled Receptors in native plasma membranes supported on beads. J Am Chem Soc 133, 16868 (2011).
[2] L Grasso, R Wyss, J Piguet, M Werner, G Hassaïne, R Hovius, H Vogel: Downscaling the analysis of complex transmembrane signaling cascades to closed attoliter volumes. Plos One (2013), in press.
[3] P Pascoal, D Kosanic, M Gjoni, H Vogel: Membrane nanotubes drawn by optical tweezers transmit electrical signals between mammalian cells over long distances. Lab on Chip 10, 2235 (2010).
Nano and sub-‐nano scale distances and dynamics from Solid State NMR in biology – an Introduction
Anthony Watts
Department of Biochemistry, University of Oxford, UK
NMR in biomolecular sciences is familiar to most, but Solid State NMR is less well represented at biophysics and structural biology meetings, despite its power to resolve very high resolution (sub-‐nanometre) distances to very high degrees of certainty (±0.05nm) and dynamics on the ns – ps time scales. As a complement to other structural biology methods, solid state NMR has a lot to offer. Here the basics of solid state NMR for use in structural biology will be presented, to provide sufficient information for participants at the main congress a background for when lectures and papers are presented in which the method is used. The nature of sample form, the way in which poorly resolved spectra using conventional solution state NMR can be enhanced, and methods for obtaining structural and dynamic data will be presented. No previous quantitative or mathematical background will be assumed, and applications will be a major focus. Further reading is given below. Suggestions for further reading: Watts, A., Straus, S.K., Grage, S., Kamihira, M., Lam, Y.-‐H. and Xhao, Z. (2004) Membrane
protein structure determination using solid state NMR. In: Methods in Molecular Biology – Techniques in Protein NMR Vol. 278 (ed. K. Downing), Humana Press, New Jersey, pp. 403-‐474.
[this chapter describes sample form, how to make various sample types, and the methodologies in some detail] Watts, A. (2005) Solid state NMR in drug design and discovery for membrane embedded
targets. Nature Reviews Drug Discovery, 4, 555-‐568. [ this review details how solid state NMR can give new insights to the dry design and discovery process. Most major pharma use solid state NMR for chemical analysis, but here the level is raised to ligand-‐target interaction studies] Levitt, M. (2008) Spin Dynamics. Wiley-‐Blackwell, Chichester. [A comprehensive NMR textbook. Very readable] Apperley, D., Harris, R., and Hodgkinson, P. (2012) Solid State NMR: Basic Principles and
Practise. Momentum Press, New York. [ a practical guide]
Submitted Abstracts (in alphabetical order of author) There are no posters. Please see programme for those selected for short oral presentations.
Symmetry and asymmetry in the unwinding of nucleic acids
Francesco Colizzi,∗,† Yaakov Levy,‡ and Giovanni Bussi∗,†
SISSA -‐ Scuola Internazionale Superiore di Studi Avanzati, via Bonomea 265, 34165 Trieste, Italy, and Department of Structural Biology, Weizmann Institute of Science, Rehovot, Israel
E-‐mail: [email protected]; [email protected]
The forming and melting of complementary base pairs in RNA and DNA duplexes are confor-‐ mational transitions required to accomplish a plethora of biological functions. Using fully atom-‐ istic simulation we have shown that RNA unwinding occurs by a stepwise process in which the probability of unbinding of the base on the 5 ́ strand is significantly higher than that on the 3 ́ strand [Colizzi and Bussi JACS, 2012]. The asymmetry in the RNA unwinding dynamics is compliant with the mechanism of helicase activity shown by prototypical DEx(H/D) RNA helicases and could allow deciphering the basis of the evolutionary pressure responsible for the unwinding mechanism catalyzed by RNA-‐duplex processing enzymes. In this spirit and from a broader standpoint, here we use a topology-‐based coarse-‐grain model to compare and characterize the mechanism of un-‐ winding for both DNA and RNA. The (a)symmetric behavior of the 3 -́‐ and 5 -́‐strand could be related to the (bi)directionality observed in molecular machineries processing nucleic acids.
∗To whom correspondence should be addressed † SISSA ‡Weizmann Institute of Science
Combining nanoscale DEER distance measurements, modelling and simulations to examine the structural heterogeneity of a peptide transporter Patricia Dijkman1, Lucy Forrest2, Philip Fowler3, Jane Kwok4, Simon Newstead4, Marcella Orwick-‐Rydmark1, Sebastian Radestock4, Firdaus Samsudin1, Nicolae Solcan4, Anthony Watts1
1Bio-‐membrane structure unit, Department of Biochemistry, University of Oxford, Oxford, UK; 2Computational Structural Biology Group, Max Planck Institute of Biophysics, Frankfurt am Main, Germany; 3Structural bioinformatics and computational biochemistry unit, Department of Biochemistry, University of Oxford, Oxford, UK; 4Department of Biochemistry, University of Oxford, Oxford, UK; Human PepT1 is a peptide transporter belonging to the major facilitator superfamily (MFS) expressed in the gastrointestinal tract where it is responsible for the uptake of dietary nitrogen through di-‐ and tri-‐peptides. It is of pharmaceutical interest as it also transports a wide-‐range of hydrophilic drugs. Crystal structures of two homologous bacterial MFS transporters, PepTSo [1] and PepTSt [2], have recently been determined in two conformational states, and a model of the outward facing state of PepTSo was generated using the repeat-‐swapping method [3]. In this study we use nanoscale distance measurements by double electron-‐electron resonance (DEER) spectroscopy, modelling and computer simulations to examine the conformational heterogeneity of PepTSo in solution. As the width of the distance distribution obtained from DEER not only contains information on the ensemble of protein conformations sampled, but also a contribution from different spin label rotamers, this combined approach allows us to disentangle these contributions and to tentatively assign DEER measurements to known conformations of the transporter. 1. Newstead S, et al. (2011) Crystal structure of a prokaryotic homologue of the mamalian
oligopeptide-‐proton symporters, PepT1 and PepT2, EMBO J 30:417 2. Solcan N, et al. (2012) Alternating access mechanism in the POT family of oligopeptide
transporters, EMBO J 31:3411 3. Radestock S and Forrest LR (2011) The alternating-‐access mechanism of MFS
transporters arises from inverted-‐topology repeats, J Mol Biol 407:698
Nanotechnology approaches to study molecular mechanisms of amyloid toxicity in Alzheimer’s disease. Elizabeth Drolle1,2, Francis Hane1, Youngjik Choi1, Brenda Lee1, Simon J. Attwood3, Arvi Rauk4, Antonin Ollagnier5, Eric Finot5 and Zoya Leonenko1,2,3.
1Department of Biology, 2Waterloo Institute for Nanotechnology, 3Department of Physics and Astronomy, University of Waterloo, Canada; 4Department of Chemistry, University of Calgary, 5Laboratoire Interdisciplinaire Carnot de Bourgogne, Universite de Bourgogne, Dijon, France. Alzheimer’s disease is a progressive neurodegenerative disease associated with amyloid fibril formation in the brain. It is now accepted that the cytotoxicity is a result of the non-‐specific interaction of toxic soluble amyloid oligomers with the surface of plasma membrane. We used atomic force microscopy (AFM), atomic force spectroscopy (AFS), frequency modulated Kelvin probe microscopy (FM-‐KPFM), Langmuir-‐Blodgett monolayer technique and surface plasmon resonance (SPR), combined with microfluidics, to study effect of membrane composition on binding of amyloid-‐β (1-‐42) peptide and fibril formation [1]. We show that cholesterol induces electrostatic domains in lipid membrane which creates a target for amyloid binding [2]. Hormone melatonin, which regulates and maintains the body's circadian rhythm, has been shown to be protective against AD, but molecular mechanism of this protection is not understood. We show that melatonin and cholesterol have the opposite effects of the lipid membrane properties which, in turn, affect amyloid binding to the lipid membrane. We used single molecule atomic force spectroscopy to study single molecule amyloid binding of A-‐beta (1-‐42) [3]. We studied the effect of amyloid inhibitor SG1 and demonstrated that this technique can be used to test inhibitor drugs for Alzheimer’s disease.
References: 1. F.Hane, E.Drolle, R.Gaikwad, E.Faught, Z.Leonenko. 2011. Journal of Alzheimer’s Disease.
26: 485-‐494; 2. E.Drolle, R.Gaikwad, Z.Leonenko. Biophysical Journal Letter, 2012, 103(4): L27-‐L29. 3. F.Hane, G.Tran, S.J. Attwood, and Z.Leonenko. 2013, PLoS ONE, 8(3): e59005.
Altering the torsional rigidity of proteins by surfactants F.A. Gutierrez1, L.J. van IJzendoorn,1 M.W.J Prins1,2 1Eindhoven University of Technology 2Philips Research Laboratories, The Netherlands Non-‐ionic surfactants are widely used in protein biosensing technologies to improve sensitivity and specificity. The surfactants inhibit protein aggregation and improve the functionalized surfaces in immunoassays. However, surfactants can also potentially alter protein conformation, ligand-‐receptor affinity and thereby assay performance. In practice, the surfactant concentration in assay buffers is an empirical compromise that is reached without design rules based on molecular understanding. Recently we have developed a torsion profiling technique [1] based on magnetic particle labels to measure the mechanical properties of individual ligand-‐receptor pairs. Using a rotating magnetic field, we apply a controlled torque to a protein pair sandwiched between a functionalized magnetic particle and a substrate, and thereby determine its torsion constant. The torsion profiling method is suited to investigate the influence of surfactants on individual ligand-‐receptor pairs in the presence of different concentrations of surfactants. Our data demonstrate an increased rotational flexibility of individual proteinG-‐IgG pairs with increasing concentration of the surfactant Tween-‐20. These results demonstrate that the mechanical integrity of the protein pair is compromised, which is most likely due to partial unfolding without breaking the ligand-‐receptor bond. Furthermore, we investigate the reversibility of the effect of the surfactant on the protein complexes. [1] A. van Reenen, F. Gutiérrez-‐Mejía, L.J. van IJzendoorn, M.W.J. Prins, Torsion Profiling of Proteins Using Magnetic Particles, Biophysical Journal 104, 1073-‐80 (2013).
Carbon nanotube as functional matrix for bacterial photosynthetic reaction centers Kata Hajdu1, Tibor Szabó1, Dóra Fejes2, Melinda Magyar1, Zsolt Szegletes3, György Váró3, Endre Horváth4, Arnaud Magrez4, Klára Hernádi2, László Forró4, László Nagy1
1Department of Medical Physics and Informatics, University of Szeged, Szeged, Hungary 2Department of Applied and Environmental Chemistry, University of Szeged, Szeged, Hungary 3Institute of Biophysics, Hungarian Academy of Science, Biological Research Center, Szeged, Hungary 4Institute of Physics of Complex Matter, Ecole Polytechnique Federale de Lausanne, Switzerland Photosynthetic reaction center protein (RC) purified from Rhodobacter sphaeroides R-‐26 purple bacterium was immobilized on –NH2 and -‐COOH functionalized and non-‐functionalized carbon nanotubes (CNTs) and the optical and electric properties of the complex was investigated. The RC binding was proved by electron microscopy and atomic force measurements. The kinetics of the absorption change after single saturating flash excitation shows that the RCs remain active in the complex for several weeks. In our experience the best activity was measured when the RC was bound physically. If the CNT/RC complex was bound to transparent conductive electrode a light induced current (photocurrent) was measured in a specially designed electrochemical cell. Light induced conductivity of the complex was also measured in a dried complex. The special electronic properties of our CNT/RC complexes open the possibility for several directions new generation applications in optoelectronics, e.g. in microelectronics or energy conversion.
Investigation of the pH stability of avidins and newly developed avidin mutants with atomic force microscopy based on single molecule sensors Melanie Koehler1, Michael Leitner1, Vesa Hytönen2, Markku Kulomaa2, Peter Hinterdorfer1, Andreas Ebner1 1Institute of Biophysics, Johannes Kepler University of Linz, Gruberstraße 40, A-‐4020 Linz, Austria 2Institute of medical Technology, Biokatu 6, FI-‐33014 University of Tampere and Tampere University Hospital, Tampere, Finland E-‐Mail: [email protected] The great stability of (strept)avidin over a wide pH range, particularly when combined with biotin, has been studied qualitatively in the last fifty years1. In the present study, a more detailed investigation and evaluation is made by performing molecular recognition studies between the receptor and their corresponding ligand, using AFM force spectroscopy. Additionally, the recently developed avidin mutant chimericavidin should be more stable against various harsh chemical conditions compared to avidin, which is also examined in this study. A high stability of the three proteins against pH treatment enables new applications in bio(nano)technology. Because of its piconewton and nanometer positional accuracy, the AFM is a powerful method for exploring the pH stability of the three proteins. The used measuring principle, so called single molecule recognition force spectroscopy, enables the investigation of forces and dynamics of the interaction between the proteins and a corresponding ligand, either on single molecule sensors, during a pH treatment and with different loading rates. Therefore, the ligand (biotin) is coupled via a hetero-‐bifunctional PEG-‐crosslinker on the outer AFM tip apex and the receptor (avidin, streptavidin or chimericavidin) is immobilized via an EGS-‐crosslinker on the probe surface. By repeatedly approaching and withdrawing of the tip in z-‐direction, receptor-‐ligand complexes are formed and released. If this experiment is repeated at different pulling speeds (loading rates) and pH values, the energy landscape of such complexes under different pHs and the pH stability itself of the receptors can be examined. The measurements have been clearly shown that the three examined proteins are stable over a wide pH range, which means that there were also protein-‐binding interactions possible at extreme pH values. Chimericavidin does not offer the pH stability on single molecule level as expected. Moreover, the energy landscape of the receptor-‐ligand complexes and the kinetic parameters during the treatment with extreme pHs could be obviously clarified. All in all, the three proteins open the possibility for more (pH-‐dependent) applications, like for e.g. in the field of pH-‐dependent regulation of protein-‐structure and/ or biotin-‐binding as well as for surface sensors, which are exposed extremes of pHs. 1NM Green. Avidin. 4. Stability at extremes of pH and dissociation into sub-‐units by guanidine hydrochloride. Biochemical Journal, 89(3):609, 1963.
Membrane-‐on-‐a-‐chip: Micro-‐structured chips to measure membrane transport and membrane fusion Ilja Kusters1, Marc Vor der Brüggen2, Sebastian Giehring2, Antoine van Oijen3 and Arnold Driessen1
1Department of Microbiology, Groningen Biomolecular Sciences and Biotechnology Institute, and Zernike Institute for Advanced Materials, University of Groningen, The Netherlands 2Nanospot GmbH, Münster, Germany 3Single Molecule Biophysics, Zernike Institute for Advanced Materials and Centre for Synthetic Biology, University of Groningen, The Netherlands Investigation of processes within biological membranes and their interfaces are hampered by the intrinsic difficulty to immobilize lipid bilayers in a functional state. For time resolved microscopic approaches, immobilization of membranes on surfaces is required but interaction of lipids or incorporated membrane proteins with the supporting surface can constrain their mobility and functionality. Here, we present the generation of free standing lipid bilayers with physiological relevant lipid composition on functionalized micro-‐structured Si/SiO2 chips that allow for both high throughput screening and single molecule imaging of the membrane. The free standing bilayers enable studies on transport processes across the membrane and fusion processes with the bilayer at physiological salt concentrations. Transport through the pore forming membrane protein hemolysin and fusion of H3N2 influenza viruses are investigated as model systems. A B
Figure 1: Free standing lipid bilayer on micro-‐structured Si/SiO2 chip. A) Micro-‐porous Si/SiO2 on a glass support suitable for high resolution fluorescence microscopy. The cavities are topped by a glass lid with 1μm apertures and a Ti/Au layer for surface functionalization. Giant unilamellar vesicles are collapsed thereby generating unilamellar lipid bilayers on top of the cavities. B) Transport through the membrane spanning pore hemolysin is monitored by efflux of small fluorophores trapped inside the chip cavity. Due to the size restriction of the pore, a larger fluorophore remains in the cavity and serves as control for membrane leakage.
Thermal properties of the S-‐layer protein from Lactobacillus salivarius Liliana Lighezan1, 3, Ralitsa Georgieva2, Adrian Neagu3
1 Faculty of Physics, West University of Timisoara, Timisoara, Romania; 2 The Stephan Angeloff Institute of Microbiology, Bulgarian Academy of Sciences, Sofia, Bulgaria; 3 Center for Modeling Biological Systems and Data Analysis, Department of Functional Sciences, Victor Babes University of Medicine and Pharmacy, Timisoara, Romania; [email protected], [email protected], [email protected] Surface layer (S-‐layer) proteins have been identified in outermost structures of the cell envelope in many organisms, such as bacteria and archaea [1]. They display intrinsic self-‐assembly property, forming monomolecular crystalline arrays [2, 3]. It is assumed that S-‐layer proteins act as protective coats, cell shape determinants, molecular and ion traps, adhesion sites for exoenzymes and structures involved in cell adhesion and surface recognition [2, 3]. Isolated S-‐layer proteins possess the unique ability to recrystallize into regular monomolecular arrays, on solid supports, on liquid surface-‐interfaces, on lipid films and liposomes, or in suspension. The ability to self-‐assemble into regular lattices, with pores of identical size and morphology (of about 1 to 10 nm), facilitates the use of S-‐layer proteins in many biotechnological applications, such as the production of isoporous ultrafiltration membranes [4] and the construction of matrices for the formation of ordered arrays of metal clusters or nanoparticles [5]. Also, they are used for drug targeting [6] and encapsulated drugs [7].
In this study, the S-‐layer protein has been isolated from Lactobacillus salivarius 16 strain of human origin, and purified by cation-‐exchange chromatography. Using circular dichroism spectroscopy, we have investigated the structure and the thermal properties of the S-‐layer protein. The far UV circular dichroism spectra indicate that the secondary structure of the S-‐layer protein consists mainly of irregular motifs, but it can also contain small fractions of α-‐helices and β-‐sheets. The near UV circular dichroism spectra show that the tertiary structure of the S-‐layer protein is determined by a high content of hydrophobic amino acids, such as Trp, Tyr and Phe, bound into a local chiral environment, which tend to compact the protein's tertiary structure. The thermal denaturation of the secondary and tertiary structures of S-‐layer protein take place in the temperature range between 40 °C and 80 °C and are partially reversible. The thermal denaturation ellipticity curves in the far and near UV domains show the existence of a metastable intermediate state in the protein denaturation pathway. During thermal denaturation, the protein changes its secondary and tertiary structure simultaneously. After the heating of the protein up to 90 °C and, subsequently, its cooling down to 10 °C, the secondary and tertiary structures of the S-‐layer protein are partially recovered, this property being important for bionanotechnological applications. References [1] Messner P and Sleytr U B, Adv. Microbiol. Physiol. 33 (1992) 213-‐275. [2] Sara M and Sleytr U B, S-‐layer proteins J. Bacteriol. 182 (2000) 859-‐868. [3] Sleytr U B and Beveridge T J, Trends Microbiol. 7 (1999) 253-‐260. [4] Neubauer A, Pum D, Sleytr U B, Biosensors and Bioelectronics 11 (1996) 317-‐325. [5] Sleytr U B, Egelseer E M, Nicola I, Pum D and Schuster B, FEBS J. 274 (2007) 323-‐334. [6] Schuster B and Sleytr U B, Curr. Nanosc. 2 (2006) 143-‐152. [7] Habibi N, Pastorino L, Soumetz F C, Sbrana F, Raiteri R, Ruggiero C, Colloids and Surfaces B: Biointerfaces 88
(2011) 366-‐372.
Engineering Biological Nanopores in Nanotechnology
Giovanni Maglia
Department of Chemistry, University of Leuven
Leuven, 3001
Ph: +32(016) 327 696 Email: [email protected]
The use of natural components as building block in nanotechnlogy is advantageous because they can be fabricated in large quantities and they can effectively interface with biological systems. However, the redesign of a protein function is difficult because its folding cannot be always predicted, and the application of proteins in nanotechnology has been limited. Proteins nanopores, which recently emerged as powerful tools in single-‐molecule investigations, are ideally suited for targeted modification because they have a stable structure and a simple geometry. Using genetic engineering, target chemical modifications and directed evolution approaches we can manipulate the size and transport properties of biological nanopores, and we can infer novel functions for desired nanotechnology applications. Examples include building a chimeric Anfinsen cage to study the chaperonian mediated protein folding at the single-‐molecule level, fabricating a selective sensor for protein analytes, and designing an artificial biological system for the vectorial transport of specific nucleic acids across biological membranes.
Photocurrent generated by photosynthetic reaction centers/carbon nanotube/ITO bio-‐nanocomposite
Melinda Magyar1 – Tibor Szabó1– Balázs Endrődi2 – Kata Hajdu1– Csaba Visy2– Zsolt Szegletes3 – György Váró3 –Endre Horváth4 – Arnaud Magrez4 – Klára Hernádi5 – László Forró4 – László Nagy1 1 Department of Medical Physics and Informatics, University of Szeged, H-‐6720 Szeged, Hungary
2 Department of Physical Chemistry and Materials Science, University of Szeged, H-‐6720 Szeged, Hungary
3 Institute of Biophysics, Hungarian Academy of Science, Biological Research Center, Szeged, Hungary 4 Institute of Physics of Complex Matter, École Polytechnique Federale de Lausanne, CH-‐1015 Lausanne, Switzerland
5 Department of Applied and Environmental Chemistry, University of Szeged, H-‐6720 Szeged, Hungary Intensive studies are focusing recently on possible directions of applications of photosynthetic proteins purified from plants (PS-‐I and PS-‐II) and from purple bacteria in nanotechnology. Different sample preparations and experimental conditions are used to find the most efficient and most stable energy converting systems. Besides the different biological systems various inorganic carrier matrices (e.g. indium tin oxide (ITO), carbon nanotubes (CNTs), silicon nanostructures) are used in different laboratories. Reaction center proteins (RC) purified from purple bacterium Rhodobacter sphaeroides were bound successfully to –NH2 and –COOH functionalized multiwalled carbon nanotubes (MWNTs) immobilized onto the surface of ITO in our studies. Electron microscopy and AFM images, flash induced absorption change and conductivity have shown that RCs can be bound effectively to the carbon nanotubes (CNT). A special electrochemical cell with three electrodes was designed for measuring the photocurrent generated by this composite. Several hundreds of nA photocurrent was measured with fully active RCs in this system which was sensitive to the conditions that fulfil conditions of the RC photo turnover.
Bacterial chemotaxis in chemical gradients created in a flow-‐free microfluidic device Krisztina Nagy, Orsolya Haja, Ádám Kerényi, Sándor Valkai, Pál Ormos, Péter Galajda Institute of Biophysics, Biological Research Centre of the Hungarian Academy of Sciences, Szeged, Hungary Motility helps bacteria explore spatially heterogeneous environments. By chemotaxis bacteria constantly detect the concentration gradient of chemoeffectors and make „decisions” on the net direction of movement (either migrate towards high attractant concentrations or away from repellents).
The technology of microfluidics is suitable for precise manipulation of liquids in microscopic dimensions in creating devices to generate stable and well-‐controlled chemical gradients. We have fabricated and experimentally characterized a microfluidic device that creates temporally stable chemical concentration gradient in a flow-‐free environment. The majority of concentration gradient generating devices are based on mixing of chemical species by the help of laminar flow, our design is a novel conception eliminating the disturbance of fluid flow. Thus bacteria in our devices swim in a static but chemically heterogeneous environment.
The device was fabricated of poly(dimethylsiloxane) using photolithography and soft lithography. It consists of two large reservoirs and a narrow observation channel between them separated by a porous membrane. Diffusion of molecules from the reservoirs to the central channel (where bacteria swim) creates the gradient across the channel. Although the gradients established in this case is less steep than in the popular flow based devices, such gradients are more than enough to observe bacterial chemotaxis.
We have studied the chemotactic response of Escherichia coli to several substances. We tested some well-‐known attractants and repellents, such as L-‐aspartate and NiSO4. We also measured the effect of „conditioned” media, cell-‐cell signaling molecules and even some antibiotics. The buffers in the reservoirs may be changed in seconds to test the cell response to an altered condition.
One of the main advantages of our device is that cells may be exposed to the gradient for extended period of time, so the behavior of the same population can be observed for long time (~ 24 h), therefore we can study small variations in chemotaxis. We are able to detect how cells change the chemical composition of their environment (consuming nutrients and releasing metabolites) and how they react to these changes.
Developing a detection system using M13 bacteriophage and Linear Dichroism
R. Pacheco-‐Gómez, S. Sandhu, M. Hicks & T. Dafforn School of Biosciences, University of Birmingham
Biomolecular detection has depended on a small number of established methodologies many of which rely on the detection of a ligand:antibody complex using an optical technique e.g. using chemiluminescence. In these systems, the ligand:antibody complex needs to be separated from bulk contaminants before a measurement can be made making the process slow, complex and expensive. This project demonstrates how a nanoparticle can provide the basis for a homogeneous assay that requires no immobilisation or separation steps. This homogeneous assay involves the use of M13 bacteriophage and linear dichroism. M13 is a filamentous bacteriophage with a high aspect ratio. This makes it easy to align in flow, allowing it to be detected using linear dichroism. This property of M13 bacteriophage has been exploited for use in a new in-‐vitro diagnostic technique that can be used to detect pathogens and could be used to detect small molecules and DNA.
Deciphering the trafficking of NK1 receptor in the membrane of living-‐cells using single-‐molecule microscopy Joachim Piguet1, Luc Veya1, and Horst Vogel1 1 Laboratory of Physical Chemistry of Polymers and Membranes, Ecole Polytechnique Fédérale de Lausanne, CH-‐1015 Lausanne, Switzerland The tachykinin receptor 1 (NK1R) is a ubiquitous 7TM receptor (GPCR) involved in
numerous functions of the nervous system, particularly in nociception, inflammation and emesis. It is a very potent target in treatment of depression and attenuation of side effects of cancer therapy. Here, we present an extensive study on the mobility distribution patterns of single receptors in the plasma membrane of living cells. In the resting state (no activation), NK1R shows two major populations: (i) freely diffusing
receptors with a narrow distribution of diffusion coefficients around 0.023 µm2/s, and (ii) receptors diffusing in confined membrane domains of 150 to 600 nm size with a broad distribution of diffusion coefficients and mobility parameters confirming significant confinement. After activating the receptor by an agonist, two successive phases occur. In the first 10 to
30 seconds after agonist binding, the fraction of freely diffusing receptors strongly decreases and a third mobility population appears, comprising immobile receptors with diffusion coefficients <10-‐3 µm2/s and mobility parameters characteristic of immobile individuals. 30 minutes later, the number of freely diffusing receptors increases again and a fourth mobility population appears comprising fast diffusing receptors in circular domains. These receptors show diffusion coefficients >0.1 µm2/s, close to ideal diffusion in lipid bilayers, and stable symmetrical domains of 300 to 600 nm size. Receptors diffusing in confined domains are directly related to clathrin-‐mediated
endocytosis (CME). Addition of drugs inhibiting different steps of CME like PitStop2, Dyngo 4a or Dynasore, leads to an accumulation of NK1R in confined regions. This membrane related event correlates with a reduction of the activity of the NK1R mediated Gαq pathway. On the other hand, depolymerisation of actin and microtubules does not significantly
modify the population of free-‐diffusing receptors, indicating absence of direct interaction of NK1R with the cytoskeleton. Nevertheless, actin depolymerisation triggers the apparition of fast diffusing receptors in circular domains even in the absence of activation. This effect is correlated with cell membrane blebs formation. Microtubule depolymerisation also significantly increases the population of receptors slowly diffusing in domains. This increase in confinement is potentially linked to inhibition of early endocytosis. In summary, the mobility pattern of NK1R in the plasma membrane of living cells is
accurately described by four modes. Depending on the receptor’s activity state and the level of modulation of both CME and cytoskeleton, the distribution of receptors in these mobility modes significantly changes in a reproducible manner. Our results point to the central importance of clathrin, not only in receptor endocytosis and turnover but also in NK1R membrane homeostasis and fine regulation of its activity.
Biophysical properties of VWF in single molecule force spectroscopy Authors: Sandra Posch1, Maria A. Brehm2, Tobias Obser2, Robert Tampé3, Reinhard Schneppenheim2, Peter Hinterdorfer1 1 Institute of Biophysics, Johannes Kepler University, Linz, Austria 2 University Medical Center Hamburg-‐Eppendorf, Department of Pediatric Hematology and Oncology, Hamburg, Germany 3 Institute of Biochemistry, Biocenter, Goethe-‐University Frankfurt, Frankfurt/Main, Germany E-‐Mail: [email protected] Von Willebrand factor (VWF) is a huge multimerizing protein playing a key role in hemostasis. VWF binds to the injured vessel wall (collagen), recruits platelets and probably leukocytes to the site of injury and binds factor VIII (important coagulation factor). Sites for collagen binding as an initial event are located in domains A1 and A3. [1] We performed Molecular Recognition Force Spectroscopy (MRFS) measurements of VWF’s specific binding domains to relevant substrates so as to classify the forces and the dynamics of these interactions. We tested several sample preparation methods. When collagen was adhered to different surfaces (MICA, uncoated glass, silicon) or bound via EGS-‐linker or Acetal-‐PEG-‐NHS-‐linker, it showed a high adhesive behavior and was therefore not usable for MRFS. However, with a sample preparation procedure using a PEG800-‐diamine layer, unspecific adhesion between tip and sample was low, most likely probably due to PEG-‐PEG repulsion. We quantified intermolecular forces, unbinding-‐length, binding probabilities, effective spring constants as well as xβ and koff between collagen III on the sample surface and rvWF A1-‐A2-‐A3-‐His bound to the tip as well as between collagen VI (sample) and rvWF A1-‐A2-‐A3-‐His (tip) and collagen VI (sample) and rvWF A1-‐A2-‐His (tip). [1] http://www.shenc.de/, 25.04.2013
Structural basis for molecular recognition of the mRNA cap by decapping DcpS enzyme determined using cap analogs with single atom modifications Anna Wypijewska1, Marius D. Surleac2, Joanna Kowalska1, Maciej Lukaszewicz1, Jacek Jemielity1,3, Martin Bisaillon4, Richard E. Davis5, Edward Darzynkiewicz1, Adina L. Milac2 and Elzbieta Bojarska1 1 Division of Biophysics, Institute of Experimental Physics, Faculty of Physics, University of Warsaw, Warsaw, Poland 2 Department of Bioinformatics and Structural Biochemistry, Institute of Biochemistry of the Romanian Academy (IBAR), Bucharest, Romania 3 Centre of New Technologies, University of Warsaw, Warsaw, Poland 4 Department of Biochemistry, University of Sherbrooke, Sherbrooke, Canada 5 Department of Biochemistry and Molecular Genetics, University of Colorado, School of Medicine, Aurora, USA The cap structure (m7GpppG) existing at the 5’ end of mRNA plays a key role in the post-‐transcriptional regulation of gene expression. It allows for specific recognition of the mRNA 5’ end by multiple proteins responsible for mRNA metabolism and protects mRNA transcripts from degradation by exonucleases. When cellular role of mRNA is accomplished, the cap is hydrolyzed by mRNA decapping DcpS or Dcp2 enzymes, involved in the 3’→5’ or 5’→3’ mRNA decay, respectively. We report the structural basis for molecular recognition of the cap by C. elegans DcpS (CeDcpS) determined by merging experimental binding affinity studies of cap analogs with single atom modifications introduced into the phosphate chain with computational modeling of CeDcpS structure and docking of representative cap analogs to CeDcpS active site. Two types of cap modifications were examined, where (1) bridging oxygen atom or (2) non-‐bridging oxygen atom was replaced by CH2/NH or S, respectively.
The modeled structure of CeDcpS-‐m7GpppG revealed binding mode of a cap similar to human DcpS (HsDcpS), well-‐known from crystal structure. In both cases, cap occupies the pocket containing the histidine triad (HIT) motif, that is narrow in the region responsible for the m7Guo-‐binding and extended in the region involved in the stabilization of the second nucleoside. Thus, methylation of m7Guo 2’O or 3’O position, increasing the steric hindrance, destabilizes DcpS-‐cap complex formation, as revealed in our experiments with ARCA-‐type cap analogs. Additionally, it aborts hydrogen bonds with proximal CeDcpS residues, conserved in HsDcpS. We found a significant differences of a cap binding with respect to the type of a modification. m7GpNHppG prone to hydrogen bond formation with CeDcpS Ser246 (conserved in HsDcpS) has association constant one order of magnitude higher compared to m7GpCH2ppG, which shifts away from the HIT triad as CH2 is void of hydrogen bonding ability. The numerous interactions of CeDcpS with the second nucleoside of a cap clarify the higher binding affinity for dinucleotide m7GpSppG D2 in comparison to mononucleotide m7GpSpp D2.
Due to the variety of potential medicinal applications of DcpS inhibitors, like in spinal muscular atrophy (SMA) treatment or in the anticancer therapy, the comprehensive structural basis for cap recognition by DcpS are of potential interest for future designing of DcpS inhibitors based on the natural cap structure. Supported by the National Centre for Research and Development (02/EuroNanoMed/2011 to E.D).
List of Participants
Speakers
Cees Dekker Delft University of Technology , Kavli Institute of Nanoscience Lorentzweg 1, Delft, 2628 CJ, The
Netherlands. [email protected]
Helmut Grubmueller Max Planck Institute for biophysical Chemistry, Am Fassberg 11, Goettingen, 37077, Germany. [email protected]
Peter Hinterdorfer Institute Biophysics Johannes Kepler University, Gruberstr. 40, Linz, 4020, Austria. [email protected]
David Klenerman University of Cambridge, Department of Chemistry, Room B52, Lensfield Road, Cambridge, CB2 1EW,
United Kingdom. [email protected]
Michael Mayer University of Michigan, Dept. of Biomedical Engineering, 1101 Beal Ave, Ann Arbor, 48109, U.S.A.. [email protected]
Dimitrios Stamou University of Copenhagen, Department of Chemistry, Universitetsparken 5, Copenhagen, 2100, Denmark. [email protected]
Horst Vogel Ecole polytechnique fédérale de Lausanne, EPFL SB ISIC LCPPM CH B3 494 (Bât. CH) Station 6, Lausanne,
CH-‐1015, Switzerland. [email protected]
Anthony Watts University of Oxford, Department of Biochemistry, South Parks Road, Oxford, OX1 3QU, United Kingdom. [email protected]
Participants
Roslin Adamson University of Oxford, Department of Biochemistry, South Parks Road, Oxford, OX1 3QU, United Kingdom. [email protected]
Mohammadreza Alizadehheidari
Chalmers University of Technology, Kemivägen 10, Gothenburg, 41296, Sweden. [email protected]
Loic Arm EPFL, ISIC-‐LCPPM/Station 6, Lausanne, 1015, Switzerland. [email protected]
Juan Bolivar University of Oxford, 32 Westbury Crescent, Oxford, OX4 3RZ, United Kingdom.
Francesco Colizzi SISSA -‐ Scuola Superiore di Studi Avanzati, Via Bonomea 265, Trieste, 34136, Italy. [email protected]
Tomás Dias INESC-‐MN, Rua Alves Redol, 9 , Lisboa, 1000-‐029, Portugal. [email protected]
Patricia Dijkman University of Oxford, South Parks Road, Oxford, OX1 3QU, United Kingdom.
Elizabeth Drolle University of Waterloo, 200 University Ave W, Waterloo, N2L 3G3, Canada. [email protected]
Ana Fernandes IST/INESC-‐MN, Azinhaga da Cidade, Torre B, 1ºC, Lisboa, 1750-‐065, Portugal. [email protected]
Mykhailo Girych V.N. Karazin Kharkiv National University, Tankopiya str., 19/2, app. 47, Kharkiv, 61022, Ukraine. [email protected]
Semire Uzun Göçmen Mustafa Kemal University Medical Faculty,, Head of Biophysics Dept. Serinyol Antakya, Hatay, 31120,
Turkey. [email protected]
Iryna Goncharova Institute of Chemical Technology, Prague, Technicka 5, Prague 6, 166 28, Czech Republic. [email protected]
Fabiola Gutierrez Eindhoven University of Technology, Den Dolech 2 MBx App. Physics Group, Eindhoven, 5612AZ, The
Netherlands. [email protected]
Kata Hajdu University of Szeged, Rerrich Bela square 1., Szeged, 6720, Hungary. [email protected]
Mathew Horrocks Deparment of Chemistry, University of Cambridge, Lensfield Road , Cambridge, CB21EW, United Kingdom. [email protected]
Peter Judge University of Oxford, New Biochemistry, South Parks Road, Oxford, OX1 3QU, United Kingdom. [email protected]
Daniel Klose University of Osnabrück, Department of Physics, Barbarastr. 7, Osnabrück, 49076, Germany. [email protected]
Melanie Koehler JKU Linz, Institute of Biophysics, AFM-‐Group, Gruberstraße 40, Linz, 4020, Austria. [email protected]
Ilja Küsters Groningen Biomolecular Sciences and Biotechnology , Nijenborgh 7, Groningen, 9747 AG, The Netherlands. [email protected]
Zoya Leonenko University of Waterloo, University Avenue West, Waterloo, N2L 3G1, Canada. [email protected]
Liliana Lighezan West University of Timisoara, Bd. Vasile Parvan nr. 4, Timisoara, 300223, Romania. [email protected]
Giovanni Maglia University of Leuven, Celestijnenlaan 200G, Leuven, 3001, Belgium.
Melinda Magyar University of Szeged, Rerrich Bela square 1., Szeged, 6720, Hungary. [email protected]
Anna Mathesz Biological Research Centre, HAS, Temesvári krt. 62. , Szeged, 6726, Hungary. [email protected]
Tim Meyer Max Planck Institute for Biophysical Chemistry, Am Fassberg 11, Goettingen, 37077, Germany. [email protected]
Hendrik Mohrmann Freie Universität Berlin, Arnimallee 14, Berlin, 14195, Germany. [email protected]
Vasyl Mykuliak Taras Shevchenko National University of Kyiv, 64, Volodymyrs'ka str., Kyiv, 1601, . [email protected]
Krisztina Nagy Biological Research Centre, HAS, Temesvári krt. 62., Szeged, 6726, Hungary. [email protected]
Ashley Nord University of Oxford, Parks Road, Oxford, OX1 3PU, United Kingdom. [email protected]
Lena Nyberg Chalmers University of Technology, Kemivägen 10, Gothenburg, 41296, Sweden. [email protected]
Elena Omarova Belozersky Inst. of Physico-‐Chemical Biology, MSU, Leninskie gory 1, bld.40, Moscow, 119992, Russia. [email protected]
Raul Pacheco-‐Gomez University of Birmingham, School of Biosciences, Edgbaston, Birmingham, B15 2TT, United Kingdom. [email protected]
Joachim Piguet EPFL, ISIC-‐LCPPM/Station 6, Lausanne, 1015, Switzerland. [email protected]
Marc-‐Philipp Pfeil University of Oxford, St Edmund Hall, Queens Lane, Oxford, OX1 4AR, United Kingdom.
Sandra Posch JKU Linz, Institute of Biophysics, AFM group, Gruberstrasse 40, Linz, 4020, Austria. [email protected]
Thierry-‐Johann Robin Université Technologique de Compiègne, Rue Roger Couttolenc CS 60319, Compiegne , 60203, France. [email protected]
Manikam Sadasivam Saravanan
Utrecht University, Padualaan 8, Utrecht, 3584CH, The Netherlands. [email protected]
Sandeep Sandhu University of Birmingham, School of Biosciences, Edgbaston, Birmingham, B15 2TT, United Kingdom. [email protected]
Oleksandr Savytskyi Institute of Molecular Biology and Genetics, NASU, 150, Zabolotnogo Str.,, Kyiv, 3680, Ukraine. [email protected]
Judit Somkuti Semmelweis University, Tuzolto utca 37-‐47., Budapest, 1094, Hungary. [email protected]
Agata Szuba B CUBE, TU Dresden, Arnoldstrasse 18, Dresden, 1307, Germany. [email protected]
Garrick Taylor University of Oxford, New Biochemistry, South Parks Road, Oxford, OX1 3QU, United Kingdom. [email protected]
Marta Urbanska B CUBE, TU Dresden, Arnoldstrasse 18, Dresden, 1307, Germany.
Rémi Veneziano Institut Charles Gerhardt UMR 5253, 15 avenue Charles Flahault BP 14491, Montpellier, 34093, France. [email protected]
Luc Veya EPFL, ISIC-‐LCPPM/Station 6, Lausanne, 1015, Switzerland. [email protected]
Anna Wypijewska Division of Biophysics, University of Warsaw, Zwirki & Wigury 93, Warsaw, 02-‐089, Poland. [email protected]
List of Participants giving Research Lectures Elizabeth Drolle
Ilja Kusters Joachim Piguet Sandra Posch Melanie Köhler Giovanni Maglia Fabiola Gutierrez
Anna Wypijewska Patricia Dijkman Kata Hajdu Krisztina Nagy Raul Pacheco-‐Gomez Francesco Colizzi Melinda Magyar Liliana Lighezan